Respirometry - Whole-animal Metabolic Rates - Respirometry Equipment

Respirometry Equipment

For open flow system, the list of equipment and parts is long compared to the components of a closed system, but the chief advantage of the open system is that it permits continuous recording of metabolic rate. The risk of hypoxia is also much less in an open system.

Pumps for air flow

• Vacuum Pump: a pump is needed to push (i.e., upstream location) or pull (i.e., downstream location) air into and through the animal chamber and respirometry flow-through system.
• Subsample pump: To pull air through the analyzers, a small, stable, reliable pump is used.

Flow meter and flow controllers

• Bubble flow meters: A simple, yet highly accurate way to measure flow rates involves timing movement of bubbles of soap film up glass tubes between marks of known volume. The glass tube is connected at the bottom (for push systems) or at the top (for pull systems) to the air stream. A small rubber pipette bulb attached at the base of the tube acts as both a reservoir and delivery system for the soap bubbles. Operation is simple. First, wet the glass surface along the path bubbles travel (e.g., press the bulb so that copious amounts of soap are pushed up the glass by the air flow) to provide a virtually friction-free surface. Second, pinch the bulb so that one clean bubble is produced. With a stopwatch in hand, record the time required for the bubble to travel between marks on the glass. Note the volume recorded on the upper mark (e.g., 125 = 125 ml), divide the volume by the time required to travel between marks and the result is the flow rate (ml/sec). These instruments can be purchased from a variety of sources, but they may also be constructed from appropriate-sized, glass volumetric pipettes.
• Acrylic flow meters : Under some circumstances of high flow rates we may use simple acrylic flow meters (0 - 2.5 liters/min) to control the flow rates through the metabolic chambers. The meters are located upstream from the metabolic chambers. The flow meters are simple to use but should be calibrated twice daily for use in the respirometry system: once before recording begins (but after the animal has been sealed inside the chamber!!) and again at the end of the recording (before the animal is removed from the chamber). Calibration must be done with a bubble flow meter because the calibration marks on the acrylic meters are only approximate. For proper calibration of flow rates remember that both barometric pressure and temperature of the air streaming through the flow meter (which we assume to be equal to room temperature) must be recorded.
• Mass flow meters: The equations required for calculating rates of oxygen consumption or carbon dioxide production assume that the flow rates into and out of the chambers are known exactly. We use mass flow meters which have the advantage of yielding flow rates independent of temperature and air pressure. Therefore, these flow rates can be considered to be corrected to standard conditions (Standard Temperature Pressure). We only measure and control flow at one location—downstream from the chamber. Therefore, we must assume that the inflow and outflow rates are identical. However, during construction of the respirometry system, flow rate must be measured at all steps, across all connections, to verify integrity of flow.
• Needle valves: Mass flow meters may be purchased with mass flow controllers which permit setting flow rates. These are expensive, however. Respirometry research often will attempt to measure more than one animal at a time, which would necessitate one chamber per animal and thus controlled flow through each chamber. An alternative and more cost-effective method to control flow would be via stainless steel or carbon steel needle valves. Needle valves plus mass flow meters provides a cost-effective means to achieve desired flow rates. The valves cost about $20.

Tubing and chambers

• Tubing and connections : Various kinds of tubing can be used to connect the components of the respirometry system to and from the animal chamber. A variety of kinds of flexible tubing may be used, depending on the characteristics of the system. Acetyl, Bev-A-Line, Kynar, nylon, Tygon tubing and connectors may be used in regions of the system where oxidizing atmospheres are low (e.g., background levels of ozone only); Teflon tubing would be recommended if there is an expectation for appreciable amounts of ozone to be present because it is inert to ozone. Teflon tubes are more costly and lack flexibility.
• Metabolic chambers: Chambers may be glass jars with rubber stoppers for lids; syringe barrels for small animals and insects; or constructed from Plexiglas. Ideally, chambers should be constructed from inert materials; for example, the acrylic plastics can absorb O₂ and may be a poor choice for respirometry with very small insects. Chambers need to be constructed in a manner that yields rapid mixing of gases within the chamber. The simplest metabolic chamber for a small vertebrate might be a glass jar with a stopper. The stoppers are fitted with two ports: short extensions of Teflon tubing are provided for line connections. Teflon tube extensions are pushed through the bulkhead and the line connection is finished by attaching a small hose clip to the base of the Teflon tube extension. Additionally, an extension to the inlet port inside the jar should be provided—this ensures that the animal's expiratory gases are not washed away by the in flow stream. The animal is sealed inside and the rubber stopper is held in place with Velcro straps. If an upstream system is used, any metabolic chamber leak will result in loss of animal air and, therefore, an underestimate of the animal's metabolic rate. When you close an animal inside a metabolic chamber, attention must be paid to the seal. To ensure tight seals before closing the lid, firmly work the stopper into the jar and make sure that it is even. Use 1-2 straps (2 are better) and pull tightly. Acrylic (Plexiglas) chambers will be constructed for some uses, but precise engineering will be needed to ensure proper seating; gaskets will help, and judicious use of tight-fitting clamps will minimize leaks.
• Scrubbing tubes: Water before and after the animal chamber must be removed. One arrangement would use a large acrylic column of Drierite (8 mesh (scale), i.e., relatively coarse) upstream (before the push pump, before the animal chamber) to dry incurrent airstream and several tubes with smaller mesh (10-20, i.e., relatively fine) Drierite to remove water after the animal chamber. To prepare a scrubbing tube, make sure there is a small amount of cotton at either end of the tube to prevent dust particles from traveling to the analyzers. Use small amounts of cotton, say around 0.005 g, just enough to keep the dust out of the tubing. Large amounts of cotton will block air flow when/if it gets damp. Pour the Drierite into the tube with a funnel, tap the tube on the bench to pack the grains tightly (to increase surface area - air + water rushes through loose Drierite, requiring frequent changes of scrubbers), and cap off with a small amount of cotton. To remove carbon dioxide] before and after the animal chamber, Ascarite II is used (Ascarite II is a registered trademark of the Arthur H. Thomas Co.). Ascarite II contains NaOH, which is caustic (so don't get any on your skin and keep away from water). A scrubbing tube is prepared by placing a small amount of cotton into the tube end, filling one-third of the way with 10-20 mesh Drierite, adding a small amount of cotton, then an additional third of the tube with the Ascarite II, another layer of cotton, followed by more Drierite and capping the tube off with another small amount of cotton. Tap the tube on the bench as each layer is added to pack the grains. Note: Driereite can be used over and over again (after heating in an oven), although indicating Drierite will lose color with repeated drying; Ascarite II is used once and will be considered a hazardous waste.

Analyzers

• Carbon dioxide analyzer: CO₂ analyzers typically use infrared-based detection methods to take advantage of the fact that CO₂ will absorb infra-red light and re-emit light at slightly longer wavelengths. The panel meter on the analyzer displays over the entire 0.01 - 10% CO₂ range and a voltage output proportional to CO₂ concentration is also generated for data recording.
• Oxygen analyzer: Oxygen analyzers suitable for respirometry use a variety of oxygen sensors, including galvanic ("ambient temperature"), paramagnetic, polarographic (Clark-type electrodes), and zirconium ("high temperature") sensors. Galvanic O₂ analyzers use a fuel cell containing an acidic electrolyte, a heavy-metal anode and a thin gas-permeable membrane. Since the partial pressure of O₂ near the anode is zero, O₂ is driven by diffusion to the anode via the membrane at a rate proportional to ambient O₂ partial pressure. The fuel cell produces a voltage linearly proportional to the O₂ partial pressure at the membrane. As long as cabinet temperature is stable, and provided that air flow across the fuel cell is stable and within range, the response will be 0.01% or better depending on supporting electronics, software, and other considerations.

Finally, a computer data acquisition and control system would be a typical addition to complete the system. Instead of a chart recorder, continuous records of oxygen consumption and or carbon dioxide production are made with the assistance of an analog to digital converter coupled to a computer. Software captures, filters, converts, and displays the signal as appropriate to the experimenter's needs. A variety of companies and individuals service the respirometry community (e.g., Sable Systems, Qubit Systems, see also Warthog Systems).